Prolonged or unaccustomed exercise leads to muscle cell membrane damage, detectable as release of the intracellular enzyme lactic acid dehydrogenase (LDH). This is correlated to excitation-induced influx of Ca2+, but it cannot be excluded that mechanical stress contributes to the damage. We here explore this question using N-benzyl-p-toluene sulfonamide (BTS), which specifically blocks muscle contraction. Extensor digitorum longus muscles were prepared from 4-wk-old rats and mounted on holders for isometric contractions. Muscles were stimulated intermittently at 40 Hz for 15–60 min or exposed to the Ca2+ ionophore A23187. Electrical stimulation increased 45Ca influx 3–5 fold. This was followed by a progressive release of LDH, which was correlated to the influx of Ca2+. BTS (50 μM) caused a 90% inhibition of contractile force but had no effect on the excitation-induced 45Ca influx. After stimulation, ATP and creatine phosphate levels were higher in BTS-treated muscles, most likely due to the cessation of ATP-utilization for cross-bridge cycling, indicating a better energy status of these muscles. No release of LDH was observed in BTS-treated muscles. However, when exposed to anoxia, electrical stimulation caused a marked increase in LDH release that was not suppressed by BTS but associated with a decrease in the content of ATP. Dynamic passive stretching caused no increase in muscle Ca2+ content and only a minor release of LDH, whereas treatment with A23187 markedly increased LDH release both in control and BTS-treated muscles. In conclusion, after isometric contractions, muscle cell membrane damage depends on Ca2+ influx and energy status and not on mechanical stress.
- N-benzyl-p-toluene sulfonamide
- muscle damage
- passive stretch
prolonged or unaccustomed exercise leads to muscle cell damage, detectable as a release of intracellular enzymes such as lactic acid dehydrogenase (LDH) and creatine kinase (CK). This is observed both in rats (23, 27, 30, 48) and in humans (29, 32, 40, 41, 46). Excitation causing isometric contractions, both in isolated single muscle fibers (13) and in intact muscles (6, 21–23, 38), is accompanied by an immediate and marked increase in the influx of Ca2+ leading to an intracellular accumulation of Ca2+ both in vitro (21, 22) and in vivo (16). If the cytosolic concentration of Ca2+ is elevated to critical levels for longer time periods, Ca2+-dependent proteases (calpains), or phospholipases may be activated. Activated calpains degrade ultrastructural filaments such as myofibrils or the cytoskeleton (4, 5), whereas phospholipases, when activated, induce damage to the sarcolemma (15). Increased cytosolic Ca2+ [Ca2+]i may also augment the production of reactive oxygen species (ROS) leading to peroxidation of membrane lipids (47), thereby contributing to damage of the cell membrane. The ensuing increase in permeability of the membrane allows passive influx of extracellular Ca2+ down its electrochemical gradient and intracellular enzymes (such as LDH and CK) to leak out. Stimulation-induced increased permeability of the membrane is evident from the demonstration of persisting and marked increase in 45Ca-uptake and release of LDH in resting isolated rat muscles following exposure to electrical stimulation (22, 23, 38).
N-benzyl-p-toluene sulfonamide (BTS) has been shown to specifically block muscle contraction in fast-twitch muscle fibers, by interfering with the cross-bridge cycling of the contractile filaments both in amphibians and in rodents (9, 44). BTS selectively blocks the ATP utilization of cross-bridge cycling without blocking the ATP utilization of the SR Ca2+ pump (54). The electrophysiological membrane properties (35) and the excitation-induced intracellular Ca2+ concentration ([Ca2+]i) transient (9, 14, 44) do not seem to be affected by BTS, whereas the active tension and stiffness are reversibly depressed to very low levels (9, 14, 44). It is proposed that BTS decreases the Ca2+ sensitivity of the contractile apparatus (44) and that it prevents active force generation by inhibiting the release of Pi in the actomyosin-mediated ATP hydrolysis pathway (49).
It has been proposed that during eccentric exercise, physical injury to the cell membrane is the initial event in muscle damage (1, 8, 52). Therefore, we were interested in determining whether mechanical stress induced by isometric contraction caused disruption of the sarcolemma. Passive lengthening of mouse extensor digitorum longus (EDL) muscles causes no decrease in the number of fibers or in the maximum tetanic force (18, 34, 37), neither does it affect [Ca2+]i (3). In line with this, in normal cultured myotubes from hamsters, no CK release was observed following stretch (25). The effects of passive stretch on cellular integrity in rat EDL muscle were also explored in the present study.
We tested the following three hypotheses: 1) During isometric contraction, the excitation-induced Ca2+ uptake is due to mechanical stress arising from the contraction. This is investigated by measuring the uptake of the isotope 45Ca, used as a tracer for Ca2+, in stimulated muscles in the absence and presence of BTS. 2) Muscle cell damage induced by isometric contractions is due to the mechanical stress imposed upon the muscle. This is investigated by selectively blocking the contraction of fast-twitch muscle with BTS, thus providing an opportunity to separate the effects of mechanical stress from those of excitation-induced Ca2+ accumulation on damaging processes in muscle cells. 3) The effects of an increased influx of Ca2+ depend on the energy status of the cell, as well as on the nature of the Ca2+ influx (physiological vs. ionophore mediated). This hypothesis is tested by measuring ATP and creatine phosphate contents in muscles exposed to electrical stimulation under oxygenated or anoxic conditions or treatment with a calcium ionophore, A23187, in the absence and presence of BTS.
All experiments were carried out using 4-wk-old female or male Wistar rats (own breed) all weighing between 60 and 70 g. Animals of this size were chosen to obtain muscles of a relatively small size to improve diffusion and oxygenation during incubation. The rats had free access to food and water and were maintained at a constant temperature (21°C) with constant day length (12 h). The animals were handled and maintained in accordance with the European Convention for the Protection of Vertebrate Animals used for Experimental and other Scientific Purposes. The animal facilities were checked by the Danish Inspectorate for Experimental Animals and the Animal Welfare Officer of the Medical Faculty of the University of Aarhus.
Muscle preparation and incubation.
Animals were killed by cervical dislocation followed by decapitation, and intact EDL muscles, all weighing between 20 and 30 mg, were excised as previously described (10). The standard incubation medium was a Krebs-Ringer bicarbonate buffer (pH 7.2–7.4) containing in mM: 122.1 NaCl, 25.1 NaHCO3, 2.8 KCl, 1.2 KH2PO4, 1.2 MgSO4, 1.3 CaCl2 and 5.0 d-glucose. In some experiments, the Ca2+ concentration was lowered to 0.1 or 0.3 mM. After preparation, all muscles (resting as well as stimulated) were mounted for isometric contractions on muscle holders with platinum wire electrodes placed on either side of the central part of the muscle. The buffer was continuously gassed with a mixture of 95% O2 and 5% CO2. All muscles were equilibrated for a minimum of 30 min in the standard medium before further treatment. This procedure has been shown to allow the maintenance of Ca2+, Na+, and K+ contents for several hours in vitro (11, 16). Incubations took place at 30°C in a volume of 5- or 8-ml buffer. This is the highest temperature at which the muscles are stable for hours in vitro (43). In experiments with BTS, muscles were incubated with BTS (50 μM) for 90 min before stimulation or the addition of A23187. This long preincubation was required to obtain full development of force inhibition (35). BTS was dissolved in DMSO; therefore, DMSO was added to the controls in an equivalent amount (0.05%). At this concentration, DMSO does not significantly alter the force development compared with controls without DMSO (35). Maximum isometric forces at 90 and at 40 Hz were measured after the equilibration period before any treatment, and the mean values for 18 muscles were 0.403 ± 0.06 N and 0.293 ± 0.06 N (16.1 N/g wet wt and 11.7 N/g wet wt, respectively).
EDL muscles were exposed to fatiguing stimulation using a protocol that has previously been shown to elicit muscle cell damage and long-lasting loss of force (38). The muscles were stimulated intermittently (10 s on, 30 s off) at 40 Hz (1-ms pulses of 10 V). Different groups of muscles were stimulated for 0, 15, 30, or 60 min.
Stimulation-induced 45Ca uptake.
During the last 15 min of fatiguing stimulation, the muscles were incubated in buffer containing 45Ca (0.5 μCi/ml). Immediately after incubation the muscles were washed 4 × 30 min at 0°C in Ca2+-free buffer containing 0.5 mM EGTA to remove extracellular 45Ca. Finally, the tendons were removed, and the muscles were blotted, weighed, and soaked overnight in 3 ml 0.3 M trichloroacetic acid (TCA). The next day 45Ca activity of the TCA extract was determined by liquid scintillation counting (Packard, Tri-Carb 2100 TR), and 45Ca uptake was calculated on the basis of this and the specific activity of 45Ca in the incubation medium. The uptake was corrected for loss of intracellular 45Ca during washout by a previously determined correction factor of 1.6 (21).
At the end of the experiment, the tendons were removed, and the muscles were blotted, weighed, and soaked overnight in 3 ml 0.3 M TCA to extract Na+, K+ and Ca2+. Previous studies showed that this procedure was as efficient in extraction of ions as homogenization and subsequent centrifugation of the TCA extract (22). Ca2+ content was determined by atomic absorption spectrophotometry (Solaar AAS, Thermo, Cambridge, England) using 1.5 ml of the TCA extract mixed with 150 μl of 0.27 M KCl. The muscle extracts were measured against a blank and standards containing 12.5 or 25 μM Ca2+, and the same amount of TCA and KCl as the muscle extracts. Na+ and K+ contents of the TCA extracts were determined using a Radiometer FLM3 flame photometer (Copenhagen, Denmark) with lithium as an internal standard. For each 0.5 ml sample of the TCA extract, 1.5 ml, 5 mM LiCl, and 0.5 ml 0.3 M TCA were added.
As an indicator of cellular integrity, the release of LDH from the muscles into the incubation medium was determined as previously described (22). In short, after being mounted on muscle holders in 5- or 8-ml chambers but before measurement, the muscles were prewashed 4 × 30 min to remove LDH released from cells damaged during excision of the muscles. Buffer samples for determination of the level of LDH release before the fatiguing stimulation were taken from the last of the prewash tubes. During recovery, after the fatiguing stimulation, the muscles were moved to new tubes every 30 min. Buffer samples were taken immediately after removal of the muscle and BSA was added to a final concentration of 0.1%. A 250-μl sample was mixed with 2.65 ml phosphate buffer (0.1 M K2HPO4 titrated with KH2PO4 to pH 7.0) containing NADH (0.3 mM) and pyruvate (0.8 mM), and the decline in the absorbance of the substrate NADH was monitored at 340 nm (Lambda 20; Perkin Elmer, Wellesley, MA) at 30°C. The activity of LDH was expressed as U·g wet wt−1·30 min−1, 1 unit (U) being the amount of enzyme that catalyzes the utilization of 1 μmol substrate/min. The total amount of LDH in EDL muscles from 4-wk-old rats was previously determined to be 585 ± 13 U/g wet wt (22).
The calcium ionophore, A23187, allows calcium to enter the cell without excitation of the muscle. Earlier studies from this laboratory showed that A23187 markedly increases 45Ca influx (threefold during 15 min) but causes no increase in resting tension (23). Thus, incubating muscle with A23187 provides the opportunity to isolate the effects of calcium overload from those of mechanical strain. Muscles were mounted on the muscle holders in 5-ml tubes containing normal KR buffer. After prewash (4 × 30 min), muscles were incubated in buffer with A23187 (2 × 10−5 M), and LDH release was measured every 30 min in the following 3 h as described above.
The experimental setup used for the application of constant passive mechanical stretch and force measurements allowed for the simultaneous recordings from four muscles in incubation chambers containing 8 ml buffer. EDL muscles were mounted on a force displacement transducer (Grass FTO3, W. Warwick, RI), and during repeated stimulation with single pulses adjusted to optimal length (baseline tension averaging 1.0 g or 9.8 × 10−3
For the experiments with dynamic passive stretch, two EDL muscles were tied together in extension of one another. Then they were mounted with one muscle being placed between the electrodes and the other one placed above. Muscle length was adjusted to estimated resting length. After stimulation, each pair of muscles was separated, and the muscles were mounted at estimated resting length individually in new tubes for the remainder of the experiment.
For the experiments with constant passive stretch, single muscles were mounted between electrodes and adjusted to optimal length for twitch force development. Following the 4 × 30 min prewash period the passive tension of the muscles were adjusted to 0.29 or 0.39 N (30 or 40 g), corresponding to the forces developed at 40 and 90 Hz in the stimulation experiments. These tensions were maintained for 180 min and were checked approximately every 10 min and adjusted if necessary. Buffer samples were taken for determination of LDH activity in both types of experiments.
In the experiment in which muscles were exposed to anoxia, the incubation buffer was changed after the equilibration period to a buffer pregassed with 95% N2 + 5% CO2. For the remainder of the experiment, the muscles were gassed with 95% N2 and 5% CO2. Muscles exposed to anoxia were either resting for 240 min or stimulated for 60 min followed by 180 min of rest. Muscles treated with BTS were preincubated with BTS for 90 min before exposure to stimulation and anoxia. BTS was furthermore present during the 60 min of stimulation but not in the following 180 min of rest.
ATP and creatine phosphate contents.
Immediately after treatment/stimulation muscles were frozen in liquid nitrogen and stored at −80°C. Muscle samples were freeze-dried, freed from connective tissue and blood, extracted with HClO4 (0.5 M), and neutralized with KHCO3 (2.2 M), as previously described (24). The neutralized extract was analyzed for ATP and creatine phosphate (CrP) by enzymatic techniques (24).
Chemicals and isotope.
All chemicals were of analytical grade. BTS and A23187 were obtained from Sigma Aldrich (St. Louis, MO); NADH and pyruvate were from Boehringer Mannheim (Mannheim, Germany). 45Ca (1.31 Ci/mmol) was obtained from Amersham International (Aylesbury, UK).
Results are given as means ± SE. The statistical significance between groups was ascertained using two-way ANOVAs, with repeated measures for the independent variable (time) and the dependent variable (treatment) for each set of analyses, with Bonferroni post hoc tests. The alpha level in all of the analyses was 0.05.
As shown in Fig. 1, control experiments, testing force development at 90 Hz every 30 min for 5 h, showed no significant drop in maximum force during this long incubation period. The statistical power in this experiment is 1.00 (n = 11).
Effects of BTS on force development and 45Ca uptake.
Figure 2 shows that in muscles exposed to 60 min of intermittent stimulation at 40 Hz, force development declined from 100% to 50% of initial force within 3 min. Within 10 min of stimulation, force reached 14% of the initial level followed by a further slow reduction to 5–6% at the end of stimulation. Muscles incubated with BTS (50 μM) for 90 min before stimulation, showed a force production of only 8% of initial force at the onset of stimulation, and after 10 min of stimulation, almost no force (2–3% of initial) was developed.
The initial stimulation-induced increase in 45Ca uptake has previously been shown to be 17-fold, as measured over the first 30 s of stimulation at 40 Hz (22). In the present experiments, we found that stimulation for 30 s (40 Hz, 10 V, 1 ms) induced a 21-fold higher uptake of 45Ca in control muscles and a 19-fold higher uptake in BTS-treated muscles compared with resting controls (Table 1). As shown in Fig. 3, intermittent stimulation (10 s on, 30 s off) at 40 Hz for 15–60 min increased total 45Ca uptake three- to fivefold when measured over 15-min periods. Assuming that the increase in 45Ca uptake only occurs during the 10-s stimulation periods, this corresponds to a 12- to 20-fold increase. In the presence of BTS (50 μM), excitation induced the same or in one instance a slightly higher increase in 45Ca influx as in the controls without BTS. Thus, reducing the mechanical stress causes no significant reduction in the excitation-induced 45Ca uptake, neither in its early phase (0–30 s) nor during longer-lasting repetitive stimulation.
Contents of ATP and CrP.
Because BTS almost completely inhibits muscle contraction, the utilization of ATP by the contractile filaments must be reduced. This may lead to a better energy status of the BTS-treated muscles. To elucidate this, the contents of ATP and CrP were measured in stimulated muscles incubated with or without BTS. As shown in Fig. 4, A and B, BTS caused no change in ATP or CrP contents in resting muscles. The ATP and CrP contents in resting oxygenated controls were ∼29 and 88 mmol/kg dry wt, respectively. When muscles were exposed to electrical stimulation, the contents of ATP and CrP dropped by 49% and 54%, respectively, in control muscles. In muscles treated with BTS, the reductions induced by electrical stimulation were only 32% for ATP and 42% for CrP. The BTS-induced changes in ATP and CrP contents of the stimulated muscles were statistically significant (P < 0.001 for ATP and P < 0.05 for CrP), indicating that BTS improves the energy status in stimulated muscles.
Effects of BTS, anoxia, and low extracellular Ca2+ concentration on excitation-induced membrane damage.
A better energy status may render the muscle better suited to handle the incoming Ca2+. Therefore, we tested whether the BTS-treated muscles, which show the same uptake of Ca2+ but have a better energy status, respond differently with regard to loss of cellular integrity, visualized by the release of LDH. Fig. 5 shows that during the 60 min of intermittent stimulation, there was no significant increase in LDH release, either in the absence or in the presence of BTS. In the following resting period, however, LDH release from control muscles increased dramatically from ∼0.4 U·g−1 wet wt−1·30 min−1 before stimulation to ∼4.5 U·g wet wt−1·30 min−1 60 min after cessation of stimulation. BTS completely suppressed this delayed and pronounced stimulation-induced LDH release. There was no significant difference between LDH release from stimulated muscles treated with BTS and that from resting control muscles.
To determine whether the protective effect of BTS on LDH release was related to the higher ATP level, we exposed muscles to complete anoxia for 4 h (stimulation for 1 h and a following recovery period of 3 h). We have earlier made use of this anoxic protocol to induce membrane damage (20). We hypothesized that anoxia would lead to a low energy status and that, despite the presence of BTS, the combined effect of excitation-induced Ca2+ influx and reduced energy status would lead to membrane damage. As shown in Fig. 6, resting muscles exposed to anoxia showed a trend toward an increase in LDH release. However, still after 4 h of anoxia, this increase was not statistically significant (P = 0.16, n = 3). In line with an earlier study (20), muscles exposed to both stimulation and anoxia showed a pronounced LDH release, reaching 6–7 U/g wet wt/30 min, and this was not prevented by BTS. After 4 h of anoxia, the ATP levels in the two groups of stimulated muscles (±BTS), were reduced to 5.6 ± 0.2 mmol/kg dry wt (−BTS, n = 3) and 4.3 ± 1.0 mmol/kg dry wt (+BTS, n = 6). These values did not differ significantly between the two groups (P = 0.4). Force measurements in anoxic muscles showed a faster decline during the fatiguing stimulation protocol compared with oxygenated control muscles (data not shown). Thus despite the fact that mechanical stress is reduced in stimulated anoxic muscles, these muscles showed a larger degree of membrane damage than oxygenated muscles.
By lowering extracellular Ca2+ concentration ([Ca2+]o) to 0.3 or 0.1 mM (Fig. 5), the maximum stimulation induced LDH release was reduced by 37 and 54%, respectively, compared with the LDH release from stimulated muscles incubated at 1.3 mM [Ca2+]. This reduction in LDH release cannot be explained by lowered maximum force development in muscles incubated at 0.1 or 0.3 mM Ca2+. As shown in Fig. 7, muscles incubated at 0.1 or 0.3 mM Ca2+ showed no significant changes in the initial maximum force development compared with those incubated at 1.3 mM Ca2+. However, when comparing the force-time integral for the three curves, the integral for 0.3 mM and 0.1 mM [Ca2+] is significantly lower (30–34%) than the integral for 1.3 mM [Ca2+] (P = 0.02 and P = 0.04, respectively). No difference was observed between the integrals for 0.1 and 0.3 mM [Ca2+]. We believe that the greatest risk of damage to the membrane would be at the highest forces and not when force was reduced to less than 10% of initial. Therefore, despite the same maximum mechanical stress on muscles incubated at varying [Ca2+]o, reducing [Ca2+]o, and thereby reducing the stimulation-induced uptake of Ca2+ (21), led to a smaller release of LDH. As shown in Fig. 8A, the importance of Ca2+ in the damaging process is further supported by a significant correlation between LDH release and the total cellular Ca2+ content of the muscles (r2 = 0.71, P = 0.001). Membrane damage is also evident from the loss of K+ and gain of Na+, as shown in Fig. 8B. LDH release shows a significant positive correlation to the loss of K+ (r2 = 0.68; P = 0.002) and a significant negative correlation to the gain of Na+ (r2 are 0.67; P = 0.002).
In the present study, the accumulated release of LDH from the muscles never exceeds 4.3% (Fig. 5) of the total content of LDH in a rat EDL muscle (585 ± 13 U/g wet wt; Ref. 22). Thus only a small fraction of the total content of LDH is released from the muscles. This is in line with studies on the increase in plasma LDH and CK after prolonged exercise in humans (42). Unfortunately, our data do not allow differentiating between widespread but partial cell damage and localized but complete loss of cellular integrity.
Effects of BTS on A23187-induced membrane damage.
The effect of Ca2+-overload without a mechanical component was tested using the Ca2+ ionophore A23187. Earlier studies from this laboratory showed that A23187 markedly increases 45Ca influx (threefold during 15 min) but causes no increase in resting tension (23). As shown in Fig. 9, A23187 markedly increased LDH release reaching 7–8 U/g wet wt/30 min after 120 min both in the absence and in the presence of BTS. Thus, Ca2+ influx alone, with no mechanical component, can cause LDH release, irrespective of the presence of BTS. This also indicates that BTS as such does not interfere with the processes of LDH release and therefore cannot explain the inhibitory effect of BTS on the excitation-induced LDH release.
To elucidate whether the LDH release in A23187-treated muscles was induced by reduced ATP and CrP contents or by Ca2+ influx alone, ATP and CrP contents were also measured in these muscles. As shown in Fig. 4, A and B, there were no changes in the contents of ATP or CrP in A23187-treated muscles compared with normal resting muscles. Furthermore, the energy status in A23187-treated muscles was not affected by BTS.
Passive mechanical stretching.
To identify a possible effect of mechanical stress alone on membrane damage, two types of experiments were performed: dynamic and constant stretch. In the first experiment, two muscles were tied together in extension of one another. The upper muscles were exposed to dynamic passive stretch when the lower muscles were actively contracting. As shown in Fig. 10A, LDH release was 14.2-fold higher in the actively contracting muscles compared with their resting controls, whereas LDH release in the muscles exposed to dynamic passive stretch was only 1.6-fold higher than in their resting controls. The total cellular Ca2+ contents were also measured at the end of this experiment. Compared with the resting control level, Ca2+ content was not increased in muscles that had been exposed to dynamic passive stretch (1.84 vs. 2.00 μmol/g wet wt, P = 0.5). In contrast, the total cellular Ca2+ content in the muscles exposed to active contraction was 94% higher (P < 0.001) than in their resting control muscles (3.71 vs. 1.92 μmol/g wet wt).
In the second experiment, muscles were exposed to constant passive stretch at 0.29 or 0.39 N for 180 min, which is comparable to the tension developed during the tetanic contractions elicited by stimulation at 40 or 90 Hz. As shown in Fig. 10B, even after 180 min of constant passive stretch of 0.29 or 0.39 N, no increase in the release of LDH was observed.
The major results of the present study are the following: 1) Excitation-induced 45Ca influx is early in onset and is unaffected by mechanical stress. 2) In muscles contracting isometrically, excitation induces a late membrane damage, which is not due to mechanical stress but depends on the excitation-induced influx of Ca2+. 3) The effects of an increased influx of Ca2+ on membrane leakage depend on the energy status of the cell, as well as on the nature of the Ca2+ uptake.
Thus, the late membrane damage following isometric contractions may occur as a result of an increased excitation-induced influx of Ca2+ in combination with a reduced energy status of the cell. In cells with a lower energy status, the Ca2+ entering from the outside may not be adequately cleared from the cytosol and stored in the SR and the mitochondria. This could cause local increases in [Ca2+]i, which may activate degradative processes within the cell leading to cell membrane damage and LDH release. Early studies by Jones et al. (27) showed that when mouse soleus muscle is stimulated electrically, LDH is released and that this LDH release is exaggerated if the muscles are stimulated under hypoxic conditions. Furthermore, LDH release from ATP-deprived mouse skeletal muscle (achieved by hypoxia, cyanide, or dinitrophenol) depends on the presence of external Ca2+ (26, 28). Studies on chick skeletal muscle have shown that protein degradation is greatly stimulated if high [Ca2+]o is combined with ATP depletion (17). These observations are in good agreement with our latest study concerning the effect of anoxia on muscle damage (20). We found that in isolated rat muscles, anoxia induces increases in Ca2+ influx and LDH release and that the degree of cell damage is very dependent on [Ca2+]o.
Excitation-induced Ca2+ uptake and cell damage.
Several studies show that excitation increases 45Ca uptake in skeletal muscle (6, 21–23). Also, in single-twitch fibers from the semitendinosus muscle of the frog, electrical stimulation causes a marked increase in 45Ca uptake (13). It should be noted that in intact rat muscle, this effect is early in onset and quite marked [up to 34-fold increase within the first 15 s of stimulation of EDL muscle (22)]. In the present study, we found a 21-fold increase during 30 s of stimulation. It has been proposed that Ca2+ influx reflects loss of cell membrane integrity brought about by increased stress at least during eccentric exercise (1). Therefore, an important motive to start the present study was that BTS offered a tool to suppress contractile force and stress without interfering with excitatory processes (35). We find that in BTS-treated muscles mounted for isometric contractions, the early phase (30 s) of excitation-induced increase in 45Ca influx is identical to that measured in the absence of BTS. Also in the BTS experiments of longer durations (15–60 min), the excitation-induced 45Ca uptake is similar to and in one instance even slightly larger than what is found in the absence of BTS (Fig. 3). This rules out that the excitation-induced stimulation of 45Ca influx is due to mechanical stress during isometric contractions. Previous studies in our laboratory (21, 38) have investigated the mechanism of excitation-induced Ca2+ influx. It was found that L-type Ca2+ channels only play a minor role in this process, and we suggested that the Ca2+ influx was mediated through Na+ channels. In the last couple of years, Ca2+ entry via store-operated Ca2+ channels (7, 31) and via channels from the transient receptor potential (TRP) family (39, 50) has been demonstrated. These channels are found widespread in most tissues in the body, and we cannot exclude that a subgroup of the TRP channels is involved in the early excitation-induced Ca2+ influx in skeletal muscles. This needs to be further investigated.
In contrast, the postexcitatory increase in the influx of Ca2+ seems to be less specific. This influx is probably due to damage to the sarcolemma. It has been suggested for many years that damage to the membrane is due to Ca2+-activated degenerative processes elicited by Ca2+ overload (4, 5, 15). The excitation-induced loss of LDH in the present study is not significant until 30 min after the cessation of a 60-min stimulation period and not fully developed until 60 min later. This argues against the idea that the LDH release is a direct consequence of increased stress. We have previously (22) and also in the present study demonstrated that the excitation-induced increase in LDH release is considerably reduced by lowering the extracellular concentration of Ca2+ from 1.3 to 0.3 or 0.1 mM (Fig. 5). This occurs despite that lowering [Ca2+]o has no effect on maximum force production during the specific fatiguing protocol used in this study (Fig. 7). In addition, the correlations shown in this paper between Ca2+ content or [Ca2+]o and LDH release (Fig. 8A) further support the hypothesis that Ca2+ plays an important role in the processes of cell damage. The cell membrane damage is also evident from the loss of K+ and gain of Na+, which were both clearly correlated to the release of LDH (Fig. 8B).
However, we also find that by suppressing contractions with BTS, the excitation-induced rise in LDH-release is abolished. The reason for this is not entirely clear. We know the difference is not due to increased permeability of the membrane of the control vs. the BTS-treated muscles, as 45Ca uptake is identical during stimulation. Therefore, BTS may affect some other parameter enabling the muscle to endure an uptake of Ca2+ that otherwise would lead to membrane damage as shown in the untreated stimulated controls. One such possible parameter could be energy status.
The electrical stimulation inducing the delayed release of LDH caused a drop in the contents of ATP and CrP of 49 and 54%, respectively. These are typical reductions observed with intense electrical stimulation in vitro (2, 36) showing that the energy production is clearly lower than the energy utilization. In muscles treated with BTS, the energy utilization for cross-bridge cycling is markedly reduced, but still, the Ca2+-ATPases are highly active and energy demanding. The Ca2+-ATPases in SR utilize 30–50% of the total ATP consumption in a normal fast-twitch muscle contraction (12, 33, 45, 51). In the presence of BTS, in which the excitation-induced increase in LDH release was abolished, the drop in ATP and CrP was significantly reduced. This higher level of ATP and CrP contents may protect the muscle against membrane damage. The increased availability of ATP may improve the clearance of Ca2+ from the cytoplasm into the SR. In an attempt to manipulate cellular ATP content, muscles were exposed to anoxia and electrical stimulation. In this situation ATP was depleted to very low levels in controls, as well as in BTS-treated muscles and LDH release was increased (Fig. 6). Under these conditions, BTS caused no suppression of the LDH release. This indicates that the inhibitory effect of BTS on LDH release seen in oxygenated muscles reflects a better energy status rather than suppression of contractions. Thus, the combination of excitation-induced Ca2+ influx and low energy status seems to be detrimental for the muscle, resulting in membrane damage and LDH release.
Earlier studies have shown that passive stretching of isolated skeletal muscles causes no injury to the muscle fibers (3, 18, 34, 37). In keeping with this, we find here that when EDL muscles are exposed to dynamic passive stretch (Fig. 10A), the amount of LDH released from these muscles is very small compared with the LDH released from actively contracting muscles (1.6-fold vs. 14.2-fold). In line with this, constant passive stretch up to 0.29 or 0.39 N, which is comparable to the tensions developed during the electrical stimulation at 40 or 90 Hz, respectively, showed no increase in the release of LDH, even after 180 min (Fig. 10B). This stress exposure by far exceeds the cumulative duration of tension (around 5 min) developed during intermittent stimulation at 40 Hz (Figs. 2 and 7). It indicates that the excitation-induced release of LDH is not a simple function of stress but likely depends on the cross-bridge cycling of the contractile filaments and the concomitant utilization of ATP.
A23187, Ca2+, and membrane integrity.
As a further investigation of the effects of an increased influx of Ca2+, the Ca2+-ionophore A23187 was used. In rat EDL muscle, A23187 induces a rapid increase in Ca2+ influx, which is not associated with any increase in resting tension (21). This indicates that global cytoplasmic Ca2+ does not reach levels high enough to activate the contractile filaments. In the absence of extracellular Ca2+, A23187 produces no LDH release, indicating that A23187 in itself causes no leakage of the plasma membrane. However, when extracellular Ca2+ was increased, release of LDH was observed showing that an influx of extracellular Ca2+ alone can cause membrane leakage (23). In the present study, A23187 gave rise to a rapid and large increase in the release of LDH despite no mechanical activity or excitation of the muscle. No decrease in ATP and CrP content was observed, which is in good agreement with an earlier study by West-Jordan et al. (53), showing increases in CK release in rat soleus muscles treated with A23187 with no change in CrP or ATP levels.
Thus, although we have shown that incubation with A23187 for 15 min induces the same influx of Ca2+ as electrical stimulation for 15 min (23), no marked use of ATP or CrP can be observed. We speculate that the distribution of the incoming Ca2+ is of importance for the development of membrane damage. In normal excitable muscle cells, Ca2+ enters the cytosol via specific channels. This is a normal physiological response, and the cell is adapted to handle the Ca2+ entering at those locations. However, if stimulation is intense or continues for a longer time, the Ca2+ handling capacity of the cell may be exceeded and cytosolic Ca2+ may reach levels high enough to cause activation of degradative processes leading to membrane damage. In contrast, A23187 induces unspecific Ca2+ influx at random sites in the membrane, in which the cell may not be adapted to handle Ca2+. This may give rise to local subsarcolemmal increases in [Ca2+]i, reaching levels high enough to activate degradative mechanisms leading to the large and rapid release of LDH that we observe. Global Ca2+ most likely is not affected, as no contracture is observed. The lack of change in CrP or ATP content might indicate that, in contrast to the stimulated muscles, Ca2+ was not cleared by the Ca2+-ATPases.
In conclusion, exercise-induced loss of muscle cell integrity during isometric contractions does not depend on mechanical stress but on Ca2+ influx and the ability of the cells to clear Ca2+ from the cytoplasm. This, in turn, depends on the energy status. In line with this, we have recently shown that anoxia increases the influx of Ca2+ in rat EDL muscles and markedly augments the excitation-induced leakage of LDH (20).
The study was supported by grants from the Danish Medical Research Council (No. 22-01-0189 and 22-02-0523), the Danish Biomembrane Research Centre, and Aarhus Universitets Forskningsfond.
We thank Vibeke Uhre, Tove Lindahl Andersen, Marianne Stürup-Johansen, and Ann-Charlotte Andersen for skilled technical assistance.
Parts of the results have been presented in a preliminary version (19).
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